Docking Complex-independent alignment of outer dynein arms with 24-nm periodicity in vitro

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1 2016. Published by The Company of Biologists Ltd. Docking Complex-independent alignment of outer dynein arms with 24-nm periodicity in vitro Toshiyuki Oda 1,2 *, Tatsuki Abe 1, Haruaki Yanagisawa 1, and Masahide Kikkawa 1 1 Department of Cell Biology and Anatomy, Graduate School of Medicine, The University of Tokyo, Hongo Bunkyo-ku, Tokyo, , Japan 2 Department of Anatomy and Structural Biology, Interdisciplinary Graduate School of Medicine and Engineering, University of Yamanashi, 1110 Shimokatoh, Chuo, Yamanashi, , Japan * Corresponding author address:toda@yamanashi.ac.jp Telephone: , FAX: Key words: cilia and flagella, axoneme, outer dynein arm, docking complex, microtubule JCS Advance Online Article. Posted on 1 March 2016

2 Summary statement Axonemal outer dynein arms can form 24-nm repeats by itself without docking complexes. The docking complex is likely to be a flexible scaffold rather than a molecular ruler. Abbreviations ODA: outer dynein arm, IDA: inner dynein arm, BCCP: biotin carboxyl carrier protein, DC: docking complex, DMT: outer doublet microtubule, N-DRC: nexin-dynein regulatory complex, OID linker: outer-inner dynein linker.

3 Abstract The docking complex (DC) is a molecular complex necessary for assembly of outer dynein arms (ODAs) on the axonemal doublet microtubules (DMTs) in cilia and flagella. The DC is hypothesized to be a 24-nm molecular ruler because ODAs align along the DMTs with 24-nm periodicity. In this study, we rigorously tested this hypothesis using structural and genetic methods. We found that the ODAs could bind to DMTs and porcine microtubules with 24-nm periodicities even in the absence of DC in vitro. Using cryo-electron tomography and structural labeling, we observed that the DC took an unexpectedly flexible conformation and did not lie along the length of DMTs. In the absence of DC, ODAs were released from the DMT at relatively low ionic strength, suggesting that DC strengthens the electrostatic interactions between the ODA and DMT. Based on these results, we concluded that the DC serves as a flexible stabilizer of the ODA rather than a molecular ruler.

4 Introduction The beating motions of cilia and flagella are driven by the outer and inner dynein arms (ODAs and IDAs, Fig. 1A). The arrangements of ODAs and IDAs on the outer doublet microtubules (DMTs) of the axonemes have highly ordered repeating structures with 24-nm and 96-nm periodicities, respectively (Fig. 1B). For IDAs, we have previously shown that a molecular ruler complex made of two proteins FAP59 and FAP172 determines the 96-nm repeats (Oda et al., 2014). The two proteins are Chlamydomonas homologues of mammalian CCDC39 and CCDC40 and form a 96-nm coiled-coil complex that governs both the repeat length and arrangements of IDAs, radial spokes, and nexin-dynein regulatory complexes. On the other hand, Takada et al. (2002) proposed that the 24-nm periodicity of ODAs is established by the docking complex (DC) of ODA via a similar mechanism. The DC is composed of three proteins, DC1, DC2, and DC3, and Chlamydomonas mutants lacking DC1 and DC2 fail to assemble ODAs in the flagella (Koutoulis et al., 1997; Takada et al., 2002; Wakabayashi et al., 2001). The recombinant protein complexes of the three subunits form a 24-nm-long complex, and the complex binds to the microtubules in a cooperative manner (Owa et al., 2014). DCs enter the flagella separately from the ODAs and are thought to provide microtubule-binding sites for ODAs (Takada, 1994; Wakabayashi et al., 2001). However, these previous studies do not provide evidence that the DC is actually required for the 24-nm periodicity of ODAs. In this study, we purified the ODAs devoid of the DC and found that the

5 ODAs were able to bind to DMTs and porcine brain microtubules with 24-nm periodicity independently of DC. We identified the precise positions of the subunits of DC using cryo-electron microscopy and structural labeling, and found that the DC was flexible and localized mostly away from the microtubules. Our results suggest that the 24-nm periodicity of ODA is determined by ODA itself.

6 Results and Discussion DC is not necessary for the 24-nm periodicity of ODA In order to determine whether the DC is required for the 24-nm periodicity of the ODA, it is necessary to purify ODA particles devoid of DCs. We fractionated the high-salt extract of Chlamydomonas axonemes using a MonoQ anion-exchange chromatography column (Goodenough et al., 1987). ODAs were separated into and particles, and the DC was eluted between the and peaks (Fig. S1A). As we detected trace amounts of DC in the fractions containing particles, we depleted the remaining DC by incubating the fractions with oda3 axonemes, which lack ODA and DC. Both ODA and DC bound to the oda3 axonemes, but only DCs were depleted because there was an excess of ODA in the sample (Fig. S1B). After three rounds of depletion, we successfully obtained ODA particles lacking detectable DCs (Fig. S1C and S1D). Note that we detected ODA using anti-oda intermediate chain 2 (IC2) antibody (D6168, Sigma-Aldrich). We incubated ODAs with oda3 axonemes in the presence and absence of DC (Fig. S1E), and reconstructed the three-dimensional (3D) structures of the DMTs using cryo-electron tomography (Fig. 1C and 1D). Surprisingly, the purified ODA particles arranged with 24-nm periodicities along the DMT irrespective of the presence of DC (Fig. 1D). The overall structures of the ODA with and without DC were not significantly different from one another, except for two small bumps on the outer surface of the ODA (Fig. 1D, arrowheads), suggesting that most of the densities of the DC were averaged out due to structural flexibility.

7 In order to determine whether any other axonemal proteins, such as CCDC103 (King and Patel-King, 2015; Panizzi et al., 2012), were required for the 24-nm periodicity of the ODA, we also incubated ODAs with cytoplasmic microtubules purified from porcine brains. The ODAs regularly bound to the porcine microtubules with 24-nm periodicity in the presence and absence of DC (Fig. 1E and 1F). These results suggest that the 24-nm periodicity of ODA does not require DC, and is presumably determined by ODA molecules themselves. Alignment of ODA along cytoplasmic microtubules with 24-nm periodicity has been reported previously (Haimo et al., 1979; Satir et al., 1981), but the extracted/purified ODAs used in these studies are likely to contain DCs. (Oda et al., 2007; Sakato and King, 2003). DC stabilized the interaction between ODA and DMT We hypothesized that DC reinforces the interaction between the ODA and DMT because the DC has higher affinity for DMTs when compared with the intermediate chains of the ODA (Ide et al., 2013; Owa et al., 2014). To test this hypothesis, we increased the ionic strength of the buffer used for resuspension of the ODA-reconstituted oda3 axonemes from 50 mm potassium acetate to 100 mm sodium chloride. We prepared cryo-samples of the axonemes in the high salt buffer and examined the binding of ODA to DMTs. We found that ODAs dissociated from the DMT in high-salt buffer in the absence of the DC (Fig. 2A and 2B, high salt). On the other hand, ODAs appeared unchanged in high-salt buffer in the presence of the DC. As ODAs were extracted from the axonemes with mm potassium/sodium chloride

8 (Pfister et al., 1982; Dean and Mitchell, 2013), the dissociation of ODAs from DMTs in 100 mm sodium chloride indicates that the interaction between the ODA and DMT is significantly weaker in the absence of DC when compared with that in the presence of DC. Lack of ODA in the DC-missing mutant strains is probably due to the reduced affinity of the ODA to DMT. DC localized away from DMT In order to visualize the interactions between the DC and ODA and between the DC and DMT, we structurally labeled DC1 and DC2, at four positions each (Fig. 3A, Table S1, Fig. S2), using the biotin-streptavidin system (Oda and Kikkawa, 2013; Oda et al., 2014), and identified their 3D positions in the axonemes (Fig. 3B, 3C). The densities of the labels on DC1 and DC2 appeared distant from the surface of DMTs, except for DC1-M216, which was located in close proximity to the microtubule-binding site of the ODA. Most of the labels on DC1 were located near the distal edge of the ODA. These results suggest that the DC makes contact with the DMT in the middle segment of DC1, and the rest of the complex binds to the outer surface of the ODA with heterogeneous conformations, rather than lying along the length of the DMT. The electrophoresis of the biotin-tagged DCs appeared odd; DC2M507 s band was doubled and the heights of the tagged proteins were varied (Figure S2A). As all the tagged DCs were properly biotinylated (Figure S2A and S2B) and expression of the tagged DCs restored motility defects of the mutants (Table S1), we suppose that insertion of the tags in the middle of the sequence affected the behavior of the proteins

9 during the electrophoresis and resulted in the irregular band patterns. Outer-inner dynein linkers are not required for ODA localization Although the 24-nm periodicity of the ODA appears to be determined by ODA itself, there must be a mechanism that keeps the 24-nm repeats in fixed positions relative to the 96-nm repeats. If the positions of the ODA s 24-nm repeats were not fixed relative to the 96-nm repeats, we could not visualize the structures of the two repeats simultaneously on an averaged subtomogram of the DMT. As the ODAs and IDAs are structurally crossbridged via the outer-inner dynein (OID) linkers (Fig. S3A, arrowhead) (Nicastro et al., 2006; Bui et al., 2009; Oda et al., 2013), we suspected that the OID linkers coordinate the relative positions between the 24-nm and 96-nm repeats. We generated a triple mutant ida1pf2pf3, which lacked all the major OID linkers, and examined the structural relationship between the 24-nm and 96-nm repeats (Fig. S3A). In contrast to our expectations, the 24-nm repeats of the ODAs were in the correct positions relative to the 96-nm repeats of the radial spokes. The mechanism for the structural coordination between the 24-nm and 96-nm repeats remains to be elucidated. Note that one of the OID linkers was not visible on the ODA-reconstituted oda3 axoneme structures regardless of the presence of DC (Fig. S3B, arrowheads). This result supports our conclusion above that OID linkers are not necessary for fixing the relative positions between ODAs and IDAs.

10 Role of the ATP-dependent microtubule binding sites of ODA As one ODA has ATP-dependent microtubule binding sites as well as ATP-independent binding regions (Haimo et al., 1979; Oda et al., 2007), we examined the effect of ATP on alignment of ODAs along microtubules. We incubated ODA and oda3 axonemes in the presence of ATP and vanadate to fix the ATP-dependent microtubule binding sites in weak-binding state (Fig. S3C) (Shimizu et al., 1983). We found sporadic binding of ODA, but alignment with 24-nm periodicity was not observed. We also incubated ODA with porcine microtubules in the presence of ATP and vanadate, but no decoration of ODA was observed (Fig. S3D). In the previous studies, ODAs sometimes align along a single microtubule with 24-nm periodicity only via the ATP-dependent binding sites and do not form cross-bridges (Haimo et al., 1979; Satir et al., 1981; Oda et al., 2007), suggesting that 24-nm periodicity of ODA does not require microtubule binding via the ATP-independent binding sites. Our results suggest that the ATP-dependent binding sites play the major role in the recruitment of ODA to DMT, and inter-molecular interaction between adjacent ODAs determines the periodical alignment. Note that even one ATP-dependent binding site would suffice for microtubule binding of ODA because it has been shown that ODAs lacking two of the three ATP-dependent binding sites can bind to the axonemes in vivo (Sakakibara et al., 1993).

11 Possible meaning in the flexibility of DC Our results suggest that DC is not necessary to establish the 24-nm periodicity of ODA along DMT. Instead, the DC is thought to work as a flexible adaptor that stabilizes the interaction between the ODA and DMT (Fig. 4). The structure of the DC was first observed as a small projection on the DMT by thin section electron microscopy of oda6 axonemes, which lack ODAs but have DCs (Takada et al., 1994). However, we were unable to detect significant DC-like densities in the averaged subtomogram of oda6 axonemes (Fig. S4A) (Bui et al., 2009). We compared the thin section electron micrographs of oda3 and oda6 axonemes. In agreement with our conclusion, we found that the shapes of the small projections of the DC on oda6 axonemes were highly variable and often invisible (Fig. S4B). It is counter-intuitive that the regularly aligned ODAs are stabilized by heterogeneous structures. We suppose that the structural flexibility allows the DC to accommodate the large conformational changes occurring in the mechano-chemical cycles of the ODAs (Oda et al., 2007; Movassagh et al., 2010; Lin et al., 2014). The large and fast movements of the dynein head domains might require a structurally flexible support, rather than a rigid base, to exert forces for ciliary and flagellar beating.

12 Materials and methods Strains and reagents C. reinhardtii wild-type cells were grown in Tris-acetate-phosphate (TAP) medium. To screen transformants, cells were grown on TAP agar supplemented with 10 g/ml paromomycin. Strains used for this study are listed in Table S1. Preparation of axonemes Chlamydomonas cells were deflagellated with dibucaine-hcl, and axonemes were collected by centrifugation (Piperno et al., 1977). Flagella were demembranated with 1% Nonidet P-40 in HMDENa/K buffer composed of 30 mm Hepes-NaOH ph 7.2, 5 mm MgCl2, 1 mm dithiothreitol, 1 mm EGTA, 50 mm NaCl or 50 mm CH3COOK. Purification and reconstitution of ODAs ODA particles were purified using a MonoQ anion exchange column (GE Healthcare). Peak fractions containing ODA and particles were combined and concentrated to final concentration of 2 mg/ml. ODA particles containing DC were purified using a UnoQ anion exchange column (Bio-rad) as described previously (Furuta et al., 2009). Peak fractions containing ODA particles were also concentrated to 2 mg/ml and used for reconstitution of DC (+) axonemes. For absorption and reconstitution of ODA onto oda3 axonemes, the concentrated solutions of ODA particles were dialyzed against HMDEK buffer overnight and incubated with 50 g of axonemes for two hours. For

13 decoration of microtubules, ODA particles in HMDEK buffer were mixed with 0.1 mg/ml porcine brain microtubules in the presence of 10 M paclitaxel for one hour. Construction of DC1 and DC2 expression vectors Fragments spanning from the start codon to immediately before the stop codon for the genes encoding DC1 and DC2 were amplified with PCR using genomic DNA from the wild-type strain and were then inserted into pic2 plasmids (Oda et al., 2015). We inserted the tag sequence corresponding to amino acids of Chlamydomonas acetyl-coa carboxylase biotin carboxyl carrier protein (BCCP) in the middle of the sequences of DC1 and DC2. Cryo-electron microscopy Labeling of BCCP tagged axonemes was carried out as described previously (Oda et al., 2014). Plunge-frozen grids were transferred to a JEM-3100FEF transmission electron microscope (JEOL) with a 914 cryo-transfer holder (Gatan Inc). Tilt series images were recorded at -180 C using a TemCam-F416 CMOS camera and EM-TOOLs program (TVIPS). The angular range was ±60 with 2.0 increments. The total electron dose was limited to 100 e - /Å 2. Images were recorded at 300 kev, with 6-9 m defocus, at a magnification of 25,700 and a pixel size of 6 Å. An in-column energy filter was used with a slit width of 20 ev.

14 Image Processing Image processing for subtomogram averaging was carried out as described previously (Oda and Kikkawa, 2013) using the IMOD software package (Kremer et al., 1996), custom Ruby-Helix scripts (Metlagel et al., 2007), and the PEET software suite (Nicastro et al., 2006). The effective resolutions were ~4.9 nm (Fig. S2C). Surface renderings were generated using UCSF Chimera (Pettersen et al., 2004). The EM maps of averaged DMT will be available at the EM Data Bank ( under the accession numbers EMD Statistical analysis In order to identify statistically significant differences, we applied a Student s t-test to compare wild-type and streptavidin-labeled axonemes as described previously (Oda and Kikkawa, 2013). The isosurface threshold values were t > 7.17, with a one-tailed probability of <0.1%.

15 Fundings This work was supported by CREST, the Japan Science and Technology Agency (to M.K.), the Kazato Research Foundation (to T.O.), the Takeda Science Foundation (to M.K. and T.O.), the Japan Society for the Promotion of Science KAKENHI Grant Number (to T.O.), and the Institute for Fermentation, Osaka (to T.O.). Competing interests The authors have no competing financial interests to declare. Author contributions T.O. conceived the project, T.O., T.A. and H.Y. conducted experiments, and T.O. and M.K. drafted the manuscript.

16 References Bui, K.H., Sakakibara, H., Movassagh, T., Oiwa, K., and Ishikawa, T. (2009) Asymmetry of inner dynein arms and inter-doublet links in Chlamydomonas flagella. J. Cell. Biol. 186: Bui, K. H., Sakakibara, H., Movassagh, T., Oiwa, K., and Ishikawa, T. (2009) Asymmetry of inner dynein arms and inter-doublet links in Chlamydomonas flagella. J. Cell. Biol. 186: Dean, A.B., and Mitchell, D.R. (2013) Chlamydomonas ODA10 is a conserved axonemal protein that plays a unique role in outer dynein arm assembly. Mol. Biol. Cell. 24: Furuta, A., Yagi, T., Yanagisawa, H.A., Higuchi, H., and Kamiya, R. (2009) Systematic comparison of in vitro motile properties between Chlamydomonas wild-type and mutant outer arm dyneins each lacking one of the three heavy chains. J. Biol. Chem. 284: Goodenough, U.W., Gebhart, B., Mermall, V., Mitchell, D.R., and Heuser, J.E. (1987) High-pressure liquid chromatography fractionation of Chlamydomonas dynein extracts and characterization of inner-arm dynein subunits. J. Mol. Biol. 194: Haimo, L.T., Telzer, B.R., and Rosenbaum, J.L. (1979) Dynein binds to and crossbridges cytoplasmic microtubules. Proc. Natl. Acad. Sci. U. S. A. 76:

17 Heuser, T., Raytchev, M., Krell, J., Porter, M. E., and Nicastro, D. (2009) The dynein regulatory complex is the nexin link and a major regulatory node in cilia and flagella. J. Cell. Biol. 187: Huang, B., Piperno, G., and Luck, D.J. (1979) Paralyzed flagella mutants of Chlamydomonas reinhardtii. Defective for axonemal doublet microtubule arms. J. Biol. Chem. 254: Huang, B., Ramanis, Z., and Luck, D.J. (1982) Suppressor mutations in Chlamydomonas reveal a regulatory mechanism for Flagellar function. Cell. 28: Ide, T., Owa, M., King, S.M., Kamiya, R., and Wakabayashi, K. (2013) Protein-protein interactions between intermediate chains and the docking complex of Chlamydomonas flagellar outer arm dynein.. FEBS. Lett. 587: Kamiya, R. (1988) Mutations at twelve independent loci result in absence of outer dynein arms in Chlamydomonas reinhardtii.. J. Cell. Biol. 107: Kamiya, R., Kurimoto, E., and Muto, E. (1991) Two types of Chlamydomonas flagellar mutants missing different components of inner-arm dynein. J. Cell. Biol. 112: King, S.M., and Patel-King, R.S. (2015) The oligomeric outer dynein arm assembly factor CCDC103 is tightly integrated within the ciliary axoneme and exhibits periodic binding to microtubules. J. Biol. Chem. 290: Koutoulis, A., Pazour, G.J., Wilkerson, C.G., Inaba, K., Sheng, H., Takada, S., and Witman, G.B. (1997) The Chlamydomonas reinhardtii ODA3 gene encodes a protein of the outer dynein arm docking complex. J. Cell. Biol. 137:

18 Kremer, J.R., Mastronarde, D.N., and McIntosh, J.R. (1996) Computer visualization of three-dimensional image data using IMOD. J. Struct. Biol. 116: Lin, J., Okada, K., Raytchev, M., Smith, M.C., and Nicastro, D. (2014) Structural mechanism of the dynein power stroke. Nat. Cell. Biol. 16: Lupas, A., Van, Dyke M., and Stock, J. (1991) Predicting coiled coils from protein sequences. Science. 252: Metlagel, Z., Kikkawa, Y. S., and Kikkawa, M. (2007) Ruby-Helix: an implementation of helical image processing based on object-oriented scripting language. J. Struc. Biol. 157: Movassagh, T., Bui, K.H., Sakakibara, H., Oiwa, K., and Ishikawa, T. (2010) Nucleotide-induced global conformational changes of flagellar dynein arms revealed by in situ analysis. Nat. Struct. Mol. Biol. 17: Nicastro, D., Schwartz, C., Pierson, J., Gaudette, R., Porter, M. E., and McIntosh, J. R. (2006) The molecular architecture of axonemes revealed by cryoelectron tomography. Science. 313: Oda, T., Hirokawa, N., and Kikkawa, M. (2007) Three-dimensional structures of the flagellar dynein-microtubule complex by cryoelectron microscopy. J. Cell. Biol. 177: Oda, T., Yagi, T., Yanagisawa, H., and Kikkawa, M. (2013) Identification of the outer-inner dynein linker as a hub controller for axonemal dynein activities.. Curr. Biol. 23:

19 Oda, T., and Kikkawa, M. (2013) Novel structural labeling method using cryo-electron tomography and biotin-streptavidin system. J. Struct. Biol. 183: Oda, T., Yagi, T., Yanagisawa, H., and Kikkawa, M. (2013) Identification of the outer-inner dynein linker as a hub controller for axonemal dynein activities.. Curr. Biol. 23: Oda, T., Yanagisawa, H., Yagi, T., and Kikkawa, M. (2014) Mechanosignaling between central apparatus and radial spokes controls axonemal dynein activity.. J. Cell. Biol. 204: Oda, T., Yanagisawa, H., Kamiya, R., and Kikkawa, M. (2014) A molecular ruler determines the repeat length in eukaryotic cilia and flagella. Science. 346: Oda, T., Yanagisawa, H., and Kikkawa, M. (2015) Detailed structural and biochemical characterization of the nexin-dynein regulatory complex. Mol. Biol. Cell. 26: Owa, M., Furuta, A., Usukura, J., Arisaka, F., King, S.M., Witman, G.B., Kamiya, R., and Wakabayashi, K. (2014) Cooperative binding of the outer arm-docking complex underlies the regular arrangement of outer arm dynein in the axoneme. Proc. Natl. Acad. Sci. U. S. A. 111: Panizzi, J.R., Becker-Heck, A., Castleman, V.H., Al-Mutairi, D.A., Liu, Y., Loges, N.T., Pathak, N., Austin-Tse, C., Sheridan, E., Schmidts, M. et al. (2012) CCDC103 mutations cause primary ciliary dyskinesia by disrupting assembly of ciliary dynein arms.. Nat. Genet. 44:

20 Pettersen, E.F., Goddard, T.D., Huang, C.C., Couch, G.S., Greenblatt, D.M., Meng, E.C., and Ferrin, T.E. (2004) UCSF Chimera--a visualization system for exploratory research and analysis. J. Comput. Chem. 25: Pfister, K. K., Fay, R. B., and Witman, G. B. (1982) Purification and polypeptide composition of dynein ATPase from Chlamydomonas flagella. Cell. motility. 2: Piperno, G., Huang, B., and Luck, D.J. (1977) Two-dimensional analysis of flagellar proteins from wild-type and paralyzed mutants of Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. U. S. A. 74: Rupp, G., and Porter, M.E. (2003) A subunit of the dynein regulatory complex in Chlamydomonas is a homologue of a growth arrest-specific gene product. J. Cell. Biol. 162: Sakato, M., and King, S.M. (2003) Calcium regulates ATP-sensitive microtubule binding by Chlamydomonas outer arm dynein. J. Biol. Chem. 278: Sakakibara, H., Takada, S., King, S.M., Witman, G.B., and Kamiya, R. (1993). A Chlamydomonas outer arm dynein mutant with a truncated beta heavy chain. J. Cell. Biol. 122: Satir, P., Wais-Steider, J., Lebduska, S., Nasr, A., and Avolio, J. (1981) The mechanochemical cycle of the dynein arm. Cell. Motil. 1: Shimizu, T., and Johnson, K.A. (1983) Presteady state kinetic analysis of vanadate-induced inhibition of the dynein ATPase. J. Biol. Chem. 258:

21 Takada, S., Wilkerson, C.G., Wakabayashi, K., Kamiya, R., and Witman, G.B. (2002) The outer dynein arm-docking complex: composition and characterization of a subunit (oda1) necessary for outer arm assembly. Mol. Biol. Cell. 13: Takada, S., and Kamiya. R. (1994) Functional reconstitution of Chlamydomonas outer dynein arms from alpha-beta and gamma subunits: requirement of a third factor. J. Cell. Biol. 126: Wakabayashi, K., Takada, S., Witman, G.B., and Kamiya, R. (2001) Transport and arrangement of the outer-dynein-arm docking complex in the flagella of Chlamydomonas mutants that lack outer dynein arms. Cell. Motil. Cytoskeleton. 48: Wirschell, M., Olbrich, H., Werner, C., Tritschler, D., Bower, R., Sale, W.S., Loges, N.T., Pennekamp, P., Lindberg, S., Stenram, U. et al. (2013) The nexin-dynein regulatory complex subunit DRC1 is essential for motile cilia function in algae and humans. Nat. Genet. 45: Yanagisawa, H.A., Mathis, G., Oda, T., Hirono, M., Richey, E.A., Ishikawa, H., Marshall, W.F., Kikkawa, M., and Qin, H. (2014) FAP20 is an inner junction protein of doublet microtubules essential for both the planar asymmetrical waveform and stability of flagella in Chlamydomonas. Mol. Biol. Cell. 25:

22 Figures

23 Figure 1 Reconstitution of the ODAs onto the oda3 axonemes in the presence and absence of the DC. (A) Cross-sectional views of the 3D structure of wild-type axoneme. (left) Tip-to-base view of the 9+2 structure. (right) Enlarged view of one of the DMTs. Directions of the views in B and in C, D are indicated. (B) A longitudinal view of wild-type axoneme. ODA, IDA, and nexin-dynein regulatory complex (N-DRC) are colored green, yellow, and blue, respectively. Arrows indicate the 24-nm and 96-nm periodicities of ODA and IDA, respectively. (C) The 3D structures of the wild-type (WT) and oda3 axonemes. ODAs align along DMT with 24-nm periodicity. Oda3 axoneme lacks both ODA and DC. (D, top and middle) The 3D structures of in vitro reconstituted ODA-oda3 axonemes with and without DCs. Regardless of the presence of DCs, ODAs aligned along the DMTs with 24-nm periodicity. Arrowheads indicate a small bump, which is visible only on the ODA with DC. Tip sides are on the left. (D, bottom) Difference map between two structures above. Densities colored in red indicate possible partial localizations of the DCs. The isosurface threshold of the difference map was set to two-sigma value. (E) Cryo-electron micrographs of in vitro reconstituted ODA-porcine microtubule (MT) complexes with and without DC. Even in the absence of DCs, ODAs cross-bridge between two microtubules with 24-nm periodicity. (F) Diffraction patterns of the images in F, showing the 24-nm layer-lines. (D, E) The MT-only panels present a micrograph and its diffraction pattern of porcine microtubules without ODA.

24 Figure 2 Effects of 100 mm sodium chloride on binding of ODA to oda3 axonemes. (A, B) In vitro reconstituted ODA-oda3 axonemes were resuspended and frozen in HMDE buffer containing 50 mm potassium acetate (low salt) or 100 mm sodium chloride (high salt). (A) Slices of un-averaged tomograms of axonemes. (top) In the low salt buffer, ODAs without DC aligned with 24-nm periodicity. (middle) In the presence of DC, the high

25 salt buffer did not affect the alignment of ODAs. (bottom) In the absence of DCs, most of ODAs were dissociated from DMTs in the high salt buffer. Red lines (left) indicate positions of the slices (right). (B) Averaged subtomograms also showed detachment of ODAs without DC from DMTs in the high salt buffer.

26

27 Figure 3 Structural labeling of DC subunits. (A) Coiled-coil structure prediction using CoilScan program (Lupas et al., 1991). Probabilities of coiled-coil formation of DC1 and DC2 are plotted along the amino acid sequences. Numbers indicate amino acids lengths and arrowheads indicate positions of BCCP tags. (B) 3D localizations of labels on DC1 and DC2. Tag densities were visualized by comparing the wild-type and labeled DMT structures. Colored densities correspond to positions of the labels indicated by arrowheads in the same color in A. (C) Model of DC architecture. Possible locations of DC1 and DC2 are shown in blue and red, respectively. Note that DC1 and DC2 can take different configurations due to the structural flexibility.

28 Figure 4 Model of ODA arrangement on DMTs. (left) In the absence of DCs, ODAs form a regular array with 24-nm periodicity. ODA complex (green) lean against the adjacent ODA and periodicity length is determined by the physical size of the ODA. (right) In the presence of DCs, the association between the ODA and DMT is stabilized by DCs (orange).